Streptomyces wadayamensis MppP Is a Pyridoxal 5 - ACS Publications

Nov 9, 2015 - Department of Chemistry and Biochemistry, University of Wisconsin Milwaukee, 3210 North Cramer Street, Milwaukee, Wisconsin. 53211 ...
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Streptomyces wadayamensis MppP Is a Pyridoxal 5′-PhosphateDependent L‑Arginine α‑Deaminase, γ‑Hydroxylase in the Enduracididine Biosynthetic Pathway Lanlan Han, Alan W. Schwabacher, Graham R. Moran, and Nicholas R. Silvaggi* Department of Chemistry and Biochemistry, University of WisconsinMilwaukee, 3210 North Cramer Street, Milwaukee, Wisconsin 53211, United States S Supporting Information *

ABSTRACT: L-Enduracididine (L-End) is a nonproteinogenic amino acid found in a number of bioactive peptides, including the antibiotics teixobactin, enduracidin, and mannopeptimycin. The potent activity of these compounds against antibiotic-resistant pathogens like MRSA and their novel mode of action have garnered considerable interest for the development of these peptides into clinically relevant antibiotics. This goal has been hampered, at least in part, by the fact that L-End is difficult to synthesize and not currently commercially available. We have begun to elucidate the biosynthetic pathway of this unusual building block. In mannopeptimycin-producing strains, like Streptomyces wadayamensis, L-End is produced from L-Arg by the action of three enzymes: MppP, MppQ, and MppR. Herein, we report the structural and functional characterization of MppP. This pyridoxal 5′-phosphate (PLP)-dependent enzyme was predicted to be a fold type I aminotransferase on the basis of sequence analysis. We show that MppP is actually the first example of a PLPdependent hydroxylase that catalyzes a reaction of L-Arg with dioxygen to yield a mixture of 2-oxo-4-hydroxy-5-guanidinovaleric acid and 2-oxo-5-guanidinovaleric acid in a 1.7:1 ratio. The structure of MppP with PLP bound to the catalytic lysine residue (Lys221) shows that, while the tertiary structure is very similar to those of the well-studied aminotransferases, there are differences in the arrangement of active site residues around the cofactor that likely account for the unusual activity of this enzyme. The structure of MppP with the substrate analogue D-Arg bound shows how the enzyme binds its substrate and indicates why D-Arg is not a substrate. On the basis of this work and previous work with MppR, we propose a plausible biosynthetic scheme for L-End.

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synthesis of cyclic peptides (e.g., ref 6), makes it desirable and feasible to produce TXB, END, and MPP analogues with increased potency and/or improved pharmacological properties. One of the major obstacles to making analogues of these complex natural products is the presence of L-End or its hydroxylated derivative, β-hydroxy-L-End. While an asymmetric synthesis that gives β-hydroxy-L-End with an acceptable enantiometic excess has been reported,7 it is a long (11 steps), low-yield procedure that begins from an advanced intermediate. The alternative strategy of purifying L-End from a hydrolysate of END or MPP would likely be difficult and inefficient because of the presence of multiple amino acids with similar physicochemical properties (e.g., the ornithine residues in END). Given the density of functional groups and the number of chiral centers in L-End, an enzymatic synthesis may prove to be more efficient and provide a sufficient supply of LEnd to support SAR studies of END, MPP, and TXB. To this end, we are seeking to understand the chemical logic and underlying mechanistic details of L-End biosynthesis.

he nonproteinogenic amino acid L-enduracididine [L-End (Scheme 1, 1)] is a component of a number of bioactive

Scheme 1

natural product compounds, including the non-ribosomally produced peptide antibiotics teixobactin (TXB), enduracidin (END), and mannopeptimycin (MPP).1−3 All three antibiotics have potent activity against antibiotic-resistant pathogens like methicillin-resistant Staphylococcus aureus (MRSA)1,4,5 and have a mode of action distinct from that of currently used antibiotics like vancomycin. Though all three, like vancomycin, bind the lipid II intermediate during cell wall biosynthesis, TXB, END, and MPP bind distinct regions of the lipid II molecule. The promise of these compounds for treating antibiotic-resistant infections, coupled with recent advances in the organic © XXXX American Chemical Society

Received: September 14, 2015 Revised: November 4, 2015

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DOI: 10.1021/acs.biochem.5b01016 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry Feeding studies in Streptomyces fungicidicus8 using radiolabeled amino acids established that the L-End in the antibiotic enduracidin originated from L-Arg. In addition, the END and MPP biosynthetic clusters have only three pairs of enzymes with a high degree of sequence identity: EndP/MppP (80%), EndQ/MppQ (68%), and EndR/MppR (75%).9,10 Disruption of EndP, EndQ, or EndR in S. f ungicidicus results in loss of enduracidin production that can be complemented by adding exogenous L-End to the culture medium.11 MppO, which has no homologue in the END biosynthetic cluster, is a non-heme iron, α-ketoglutarate-dependent oxygenase that selectively hydroxylates the β-carbon of free L-End to produce βhydroxy-L-End; L-Arg is not a substrate for this enzyme.12 Thus, the trio of enzymes, End/MppP, MppQ, and MppR, likely converts L-Arg to L-End. Sequence analysis suggests that End/MppP and MppQ are both fold type I PLP-dependent aminotransferases (ATases), though the EndP and MppP proteins from S. f ungicidicus and Streptomyces hygroscopicus are both ∼100 residues shorter than typical ATases. S. hygroscopicus MppR shares ∼20% sequence identity with the acetoacetate decarboxylases from Clostridium acetobutylicum and Chromobacterium violaceum. Our earlier work on this enzyme, however, showed that it is not a decarboxylase and that, in its crystalline form, it is able to cyclize 2-oxo-4-hydroxy-5-guanidinovaleric acid (Scheme 1, 2) to produce the iminoimidazolidine ring of L-End.13 There was not sufficient 2 to demonstrate catalytic turnover, but hydrolysis of the Schiff base complex captured in the crystal structure [Protein Data Bank (PDB) entry 4JME] would yield the ketone form of L-End (3). Assuming it was released from the enzyme, 3 would likely be a substrate for MppQ, which is predicted to be a fold type I PLP-dependent ATase. We have evidence that MppQ catalyzes an efficient amino transfer reaction between 2-oxo-5-guanidinovaleric acid (4) and Lalanine (manuscript in preparation). Transamination of 3 by MppQ would yield L-End. The question, then, is how the highly oxidized L-Arg derivative 2 is produced. This is a particularly interesting question, as no iron- or flavin-dependent oxygenase has been implicated in L-End biosynthesis. Herein, we report the steady state kinetic characterization and crystallographic structure of the MppP homologue from Streptomyces wadayamensis (SwMppP), a known mannopeptimycin producer.14 The structural data show that this enzyme closely resembles fold type I PLP-dependent ATases such as Escherichia coli aspartate ATase, though the SwMppP active site is subtly different. Surprisingly, our steady state kinetic and nuclear magnetic resonance (NMR) data show that SwMppP reacts with L-Arg and molecular oxygen to yield 2 directly, indicating that SwMppP catalyzes the first step in L-End biosynthesis.

TCAAGGTATGAACCGCATTCTGCCGAAAGTC-3′ (forward) and 5′-GCTCTAGATTAGCCTTGACGTTCAGCGGTATC-3′ (reverse)]. The His6-tagged SUMO-SwMppP fusion protein was expressed from E. coli BL21 Star (DE3) cells (Invitrogen Inc., Carlsbad, CA) carrying the pE-SUMOSwMppP plasmid. Cultures were grown in Luria-Bertani medium with 50 μg/mL kanamycin at 37 °C. When the cultures reached an OD600 of ∼1.0, protein expression was induced with 0.4 mM isopropyl β-D-1-thiogalactopyranoside. The temperature was reduced to 25 °C, and the cultures were grown overnight while being shaken at 250 rpm. Cells were harvested by centrifugation, resuspended in 5 mL/g of buffer A [25 mM TRIS (pH 8.0), 300 mM NaCl, 10 mM imidazole, and 100 μM PLP] supplemented with 1 mg/mL hen egg lysozyme (Hampton Research) and 0.1 mg/mL DNase I (Worthington Biochemical Corp., Lakewood, NJ). Cells were lysed using a Branson Sonifier S-450 cell disruptor (Branson Ultrasonics Corp., Danbury, CT) for sonication for a total of 10 min at 70% amplitude with 30 s pulses separated by 45 s rest periods. The temperature was maintained at or below 4 °C by suspending the steel beaker in an ice bath directly over a spinning stir bar. The lysate was clarified by centrifugation at 39000g for 45 min and then applied to a 5 mL HisTrap column (GE Lifesciences, Piscataway, NJ) at a flow rate of 5 mL/min to isolate the His6SUMO-SwMppP fusion protein. The protein was eluted by a four-step gradient of buffer B [25 mM TRIS (pH 8.0), 300 mM NaCl, and 250 mM imidazole; 5, 15, 50, and 100%]. The His6SUMO-SwMppP fusion protein eluted in the third and fourth steps and was ∼90% pure, as judged on Coomassie-stained sodium dodecyl sulfate−polyacrylamide gel electrophoresis gels. Peak fractions were pooled and dialyzed overnight against 3.5 L of 25 mM TRIS (pH 8.0), 150 mM NaCl, 100 μM PLP, and ∼3 μM SUMO protease (LifeSensors Inc.). The dialysate was passed through the HisTrap column a second time to remove the cleaved His6-SUMO tag as well as the protease. The resulting SwMppP preparation was >95% pure. The protein was desalted using a HiTrap desalting column (GE Lifesciences) into 10.0 mM MOPS (pH 6.7) and 20 μM PLP and stored at −80 °C. Selenomethionine-labeled SgMppP was expressed from T7 Express Crystal cells (New England Biolabs, Ipswich, MA) carrying the pE-SUMO-SgMppP plasmid. Cells were grown in SelenoMethionine Medium Complete (Molecular Dimensions, Newmarket, Suffolk, U.K.) with 50 μg/mL kanamycin. SgMppP was expressed and purified using the same protocols described for SwMppP. Steady State Enzyme Kinetics. All 1.0 mL kinetic assays were conducted in triplicate at 25 °C in 100 mM BIS-TRIS propane (pH 9.0). The initial velocity of SwMppP-catalyzed hydroxylation of L-Arg was measured directly by monitoring the decrease in the dioxygen concentration using a dioxygen electrode (Hansatech Instruments, Norfolk, U.K.). The concentration of L-Arg was varied from 10 to 500 μM. The kcat and Km values were determined from the initial velocity data using the equation v0 = Vm[A]/(KM + [A]), where [A] is the concentration of L-Arg, v0 is the initial velocity, Vm is the maximal velocity, and KM is the Michaelis constant. NMR Analysis of SwMppP Reaction Products. A reaction mixture (4.0 mL) containing 10.8 μM SwMppP and 2.0 mM L-Arg in 20 mM sodium phosphate buffer (pH 8.4, H2O) was incubated in a 50 mL conical vial while being stirred at 200 rpm and room temperature for 2 h. The sample was then evaporated to dryness in a CentriFan centrifugal evaporator

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MATERIALS AND METHODS Cloning, Expression, and Purification of S. wadayamensis and Streptomyces globisporus MppP. The coding sequences of SwMppP (GenBank accession code KDR62041) and SgMppP (GenBank accession code IH67_RS0105915) were optimized for expression in E. coli and synthesized by GenScript Inc. (Piscataway, NJ). The synthetic genes were both subcloned into the pE-SUMOkan expression vector (LifeSensors Inc., Malvern, PA) using primers containing BsaI and XbaI restriction sites [SwMppP, 5′-GGTCTCAAGGTATGACGACGCAACCGC-3′ (forward) and 5′-GCTCTAGATCATTAGCGGGTTTCCAGGAC-3′ (reverse); SgMppP, 5′-GCGGTCB

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Biochemistry Table 1. Crystallographic Data Collection and Refinement Statistics resolution (Å) (last shell)a wavelength (Å) no. of reflections observed unique completeness (%)a Rmerge (%)a,b multiplicity ⟨I/σ(I)⟩a no. of reflections in the working set no. of reflections in the test set Rcryst (Rfree) no. of residues no. of solvent atoms no. of TLS groups average B factor (Å2)c protein atoms ligands solvent root-mean-square deviation bond lengths (Å) bond angles (deg) coordinate error (Å) Ramachandran statistics (favored/allowed/outliers) (%)

SwMppP

SwMppP·D-Arg

41.45−2.10 (2.14−2.10) 0.97852

44.53−2.25 (2.29−2.25) 0.97856

1532933 (60186) 106188 (5236) 100.0 (100.0) 0.106 (0.734) 14.4 (11.5) 30.7 (5.2) Model Refinement 100876 5256 0.148 (0.177) 1417 903 29

256371 (12494) 79816 (3984) 90.2 (91.2) 0.106 (0.726) 3.2 (3.1) 10.0 (1.9) 76747 3015 0.162 (0.197) 1404 693 33

32.0 30.9d 35.6

34.1 43.9d 36.6

0.013 1.395 0.17 98.3/1.7/0

0.015 1.565 0.22 98.4/1.6/0

a Values in parentheses apply to the high-resolution shell indicated in the resolution row. bR = ∑(||Fobs| − scale × |Fcalc||)/∑|Fobs|. cIsotropic equivalent B factors, including the contribution from TLS refinement. dIn the unliganded SwMppP structure, “ligands” refers to the bound Cl ions, while in the D-Arg complex structure, it refers to the D-Arg-PLP unit.

overnight, crystals were treated with glycerol and flash-cooled. X-ray diffraction data for SeMet SgMppP and native SwMppP were collected at beamline 21-ID-D of the Life Science Collaborative Access Team (LS-CAT) at the Advanced Photon Source (APS). The SwMppP·D-Arg data set was collected at LS-CAT beamline 21-ID-G. Data were processed with HKL2000.15 The structure of SgMppP was determined by the singlewavelength anomalous diffraction (SAD) method using 3.5 Å resolution data collected from a crystal of SeMet-substituted SgMppP at 0.97889 Å, 61.0 eV below the tabulated K-edge wavelength for Se (0.97950 Å). AutoSHARP16 was used to determine the Se substructure, which contained 14 of the 16 Se atoms in the asymmetric unit, and calculate density-modified electron density maps. An initial model comprising ∼90% of the asymmetric unit content was built using BUCCANEER.17 Chain A of this partially refined model (with B factors set to 20.0 Å2) was used as the search model for molecular replacement in PHASER18 to phase the unliganded SwMppP data set. The molecular replacement solution was subjected to iterative cycles of manual model building in COOT19 and maximum likelihood-based refinement using the PHENIX package (phenix.refine20). Ordered solvent molecules were added automatically in phenix.refine and culled manually in COOT. Hydrogen atoms were added to the model using phenix.reduce21 and were included in the later stages of refinement to improve the stereochemistry of the model. Positions of H atoms were refined using the riding model with a global B factor. Regions of the model for translation-librationscrew (TLS) refinement were identified using phenix.find_tls_groups (P. V. Afonine, unpublished work),

(KD Scientific, Holliston, MA) at 40 °C, and the residue was resuspended in 500 μL of D2O. Control samples of 2.0 mM LArg without enzyme and 20 mM phosphate buffer alone were prepared in parallel with the reaction mixture. For all samples, one-dimensional (1D) 1H NMR spectra were recorded at room temperature on a Bruker DRX500 500 MHz spectrometer equipped with a BBI probe. To identify the structures of any reaction products present, 1H COSY, 1H−13C HSQC, and 1 H−13C HMBC spectra were recorded for the SwMppP reaction mixture. Full spectra are provided in the Supporting Information. Crystallization, Structure Determination, and Model Refinement. Initial crystallization conditions were identified by screening 15 mg/mL SwMppP against the Index HT screen (Hampton Research). After optimization, diffraction-quality crystals were obtained by the hanging drop vapor diffusion method from 30% polyethylene glycol monomethyl ether 550 (PEG MME 550), 50 mM MgCl2, and 0.1 M HEPES (pH 8.0). Drops contained 1−2 μL of protein solution at 15 mg/mL and 1 μL of crystallization solution. Crystals formed as plates or rods after 3−4 days and grew to maximal dimensions of ∼200 μm × 50 μm × 5 μm. Crystals of selenomethionine (SeMet)substituted SgMppP were grown from 10% PEG MME 5000, 5% Tacsimate, and 0.1 M HEPES (pH 7.5). SgMppP crystallized as small rods (∼100 μm × 10 μm × 10 μm). Both crystals were cryo-protected by sequential soaks in holding solutions containing 5, 10, or 20% glycerol and flashcooled by being plunged into liquid nitrogen. The structure of SwMppP with D-Arg bound was obtained by transferring crystals into 30 μL drops of the crystallization solution supplemented with 35 mM D-Arg. After being soaked C

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Biochemistry and the TLS parameters were refined in phenix.refine. Once the refinement converged (R = 0.136, and Rfree = 0.162), the model was validated using the tools implemented in COOT and PHENIX.22,23 Sections of the backbone with missing or uninterpretable electron density were not included in the final model. Side chains with poor or missing electron density were modeled in favored rotameric conformations, and the B factors were allowed to refine without additional restraints. This was done to alert end users, who are more likely to display a model colored by B factors than to pay attention to atom occupancy values or download electron density maps, that these residues are not well-defined in the electron density. The final, refined model of SwMppP, stripped of water molecules and H atoms, and with all B factors set to 20.0 Å2, was used to determine the structure of the enzyme with D-Arg bound by difference Fourier. A similar refinement protocol was used for the SwMppP·D-Arg model. Restraints for the D-ArgPLP external aldimine were generated with phenix.elbow.24 Data collection and model refinement statistics are listed in Table 1. Coordinates and structure factors for both SwMppP models have been deposited in the Protein Data Bank as entries 5DJ1 and 5DJ3.

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RESULTS AND DISCUSSION SwMppP Catalyzes the Hydroxylation of L-Arg. The S. wadayamensis homologue of MppP was cloned and overexpressed from E. coli, yielding approximately 100 mg of protein/ L of culture. Purification required the addition of pyridoxal 5′phosphate to the lysis buffer, and any time the protein was dialyzed or desalted, but PLP could be omitted from the Ni affinity chromatography buffers without issue. As purified, the protein had a pronounced greenish-yellow color. The UV− visible spectrum has a peak at 415−420 nm associated with the internal aldimine form of fold type I PLP-dependent ATases (dashed line, Figure 1A). In high-pH buffers, such as sodium phosphate (pH 8.4) or BIS-TRIS propane (pH 9.1), the enzyme loses both its color and the peak at 415 nm (t1/2 ≈ 30 min), though there is little to no visible precipitate in the solution. While there is no discrete peak for free PLP, the 415 nm peak does shift to the blue (∼397 nm) as its intensity decreases. In buffers with pH values of 0.24, well above the threshold that implies significant structural similarity.44 Among these were three proteins of known function: cystalysin from Treponema denticola (TdCSL, PDB entry 1C7N;45 11% identical, 2.1 Å rmsd), the β-cystathionase MalY from E. coli (EcMalY, PDB entry 1D2F;46 17% identical, 2.3 Å rmsd), and the glutamine aminotransferase from Thermus thermophilus HB8 (TtGln AT, PDB entry 1V2D;47 17% identical, 2.4 Å rmsd). TdCSL and EcMalY both catalyze βelimination of cysteine or cystathionine to give pyruvate, ammonia, and a sulfur-containing compound,48−50 while TtGln AT has been shown to catalyze amino transfer between a number of aromatic amino acids and α-ketoglutarate.51 All three of these enzymes are classified into subgroup 1b of the fold type I ATases (EcAAT belongs to subgroup 1a), indicating that SwMppP likely belongs in subgroup 1b, as well. The accumulation of a species absorbing at 510 nm during the reaction of SwMppP with L -Arg, likely a quinonoid intermediate, is telling, because TdCSL and a similar enzyme from Streptomyces anginosus have been shown to stabilize a quinonoid intermediate.29,52 In TdCSL and EcMalY, a tyrosine residue (Tyr64 in TdCSL) occupies the position of His29 in SwMppP. Analysis of a Tyr64Ala variant of TdCSL showed that this residue is likely involved in the nucleophilic attack on C4′ of the PLP in formation of the external aldimine, as well as in the formation of the α-aminoacrylate intermediate.53 In wildtype TdCSL, pyruvate release is rate-limiting, but in the Tyr64Ala variant, aminoacrylate formation became the ratelimiting step. Though SwMppP is closest in terms of sequence and structure to ATases and CS-lyases, it is functionally more similar to DOPA/aromatic L-amino acid decarboxylases. These enzymes stabilize a quinonoid intermediate that then reacts directly with molecular oxygen.28 Comparison of the SwMppP structure to those of the decarboxylases (PDB entries 3RCH54 G

DOI: 10.1021/acs.biochem.5b01016 Biochemistry XXXX, XXX, XXX−XXX

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Biochemistry

Figure 6. D-Arg reacts with SwMppP to form the external aldimine and becomes trapped at that point for lack of a general base to abstract the αproton. (A) Stereoview of the SwMppP active site showing D-Arg covalently bound to the PLP cofactor as the external aldimine. The chloride ion seen in the holoenzyme structure has been displaced by the α-carboxylate of D-Arg, and the guanidinium group of the substrate analogue is clamped between two aspartic acid side chains. As in Figure 4, the 2|Fo| − |Fc| electron density map contoured at 1.0σ is shown as magenta mesh and the 2|Fo| − |Fc| composite omit map, also contoured at 1.0σ, is shown as green mesh. (B) Schematic view of the active site with D-Arg bound showing potential hydrogen bonding interactions as green, dashed lines. Water molecules are shown as blue spheres. Panel A was generated using POVSCRIPT and POV-Ray. Panel B was created using a combination of MarvinSketch (http://www.ChemAxon.com) and Adobe Illustration CC 2015.

Figure 7. There are subtle changes in the active site upon formation of the external aldimine, as shown in this overlay of the holoenzyme and D-Argbound structures. The holoenzyme is colored light blue and the D-Arg-bound enzyme dark blue. Figure rendered using POVSCRIPT+ and POVRay.

H

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Biochemistry and 1JS655) shows that both decarboxylases are missing the Tyr residue that interacts with the C3′ hydroxyl of PLP (Figure 8).

the vicinity of the catalytic lysine, especially His29 of SwMppP, plays an important part in directing the chemistry of the cofactor. Analysis of SwMppP variants with mutations at Asp218 and/or His29 should illuminate this point. Hypothetical Reaction Mechanism. Our preliminary investigation of SwMppP, while solidly identifying the overall function of the enzyme, raises a number of fundamental mechanistic questions. To provide a framework for experiments designed to answer these questions, we have developed the hypothetical mechanism shown in Scheme 2. The transitions from the internal aldimine (I) to the external aldimine (II) and the first quinonoid intermediate (III, plausibly the 510 nm intermediate) are proposed on the basis of analogy to other PLP-dependent enzymes. We propose that electron-rich species III is capable of one-electron chemistry with dioxygen. In Scheme 2, we present this as formation of superoxide anion and a carbon-centered radical (IV), but there are other possibilities. After intersystem crossing (ISC), superoxide could remove a hydrogen atom from the β-carbon to give the α−β unsaturated intermediate (V) and the hydroperoxide anion. The hydroperoxide anion could then remove a hydrogen atom from V to generate the β−γ unsaturated intermediate (VI, plausibly the 560 nm intermediate) and H2O2. Hydrolysis of intermediate V would produce the 2-ketoarginine observed by NMR. Oxidation of the second quinonoid intermediate (VI) could take place in various ways. Here we show attack on H2O2 to give the hydroxylated intermediate (VII) and water, but reaction with a second molecule of dioxygen is also plausible, with the resulting hydroxyl oxygen derived from either O2 or H2O2. Finally, hydrolysis of the external aldimine VII followed by attack of Lys221 would yield 4-hydroxy-2-ketoarginine and regenerate the internal aldimine (I). The red, dotted boxes in Scheme 2 are meant to underscore the fact that these intermediates are hypothetical and that the actual mechanism remains to be determined. For example, it is also possible that the hydroxyl group of the product comes from water, as would happen if the second quinonoid (VI) reacted with an additional molecule of dioxygen that then reacted with water. This question would, of course, be answered by isotopic labeling studies together with determination of the stoichiometry of the reaction with respect to O2; both of these experiments will be performed in future work.

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Figure 8. (A) Overlaying the active sites of the internal aldimine forms of SwMppP (light blue) and the human aromatic L-amino acid decarboxylase (PDB entry 3RCH, gray) shows that despite the overall folds being quite different, the two active sites are similar in terms of the residues arrayed around the cofactor. (B) Overlaying the active sites of the D-Arg-bound SwMppP (dark blue) and carbidopa-inhibited porcine DOPA decarboxylase (PDB entry 1JS3, gray) shows that the same rearrangement of the hydrogen bonding network of D218/N300 occurs upon formation of the external aldimine. Figure rendered using POVSCRIPT+ and POV-Ray.

CONCLUSIONS The steady state data and product analysis presented here establish SwMppP as the first known PLP-dependent hydroxylase. The observation of absorbance maxima at 510 and 560 nm suggests that the reaction of SwMppP with L-Arg and dioxygen involves two quinonoid intermediates. The first quinonoid (510 nm) must be stabilized to some extent by the enzyme, because it is quite stable in the absence of O2. In this respect, SwMppP is functionally similar to the PLP-dependent DOPA/aromatic L-amino acid decarboxylases.56 SwMppP produces two products, 2-oxo-4-hydroxy-5-guanidinovaleric acid and 2-oxo-5-guanidinovaleric acid, in a ratio of ∼1.7:1. Future work will investigate whether this ratio changes with the concentration of L-Arg or O2, buffer conditions (pH and ionic strength), and/or the presence of the other L-End biosynthetic proteins. As suggested by sequence similarity, the overall fold of the enzyme is most similar to that of typical fold type I ATases. However, the structure of the internal aldimine form of the enzyme also shows that the fine structure of the active site is also more similar to that of the CS-lyases and amino acid

Also, both have an asparagine residue in place of one of the phosphate-binding serine residues of the ATases. This residue, Asn300, occupies the same position as Asp218 in SwMppP and, like Asp218, makes a hydrogen bonding interaction with the catalytic lysine side chain when an inhibitor is present [PDB entry 1JS355 (Figure 8)]. The decarboxylases have no counterpart to the unusual His29 of SwMppP, that approximate position in the decarboxylases being occupied by Thr82. However, the decarboxylases have a tyrosine (Tyr79) ∼3.0 Å away (Figure 8). It may be that the constellation of residues in I

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Biochemistry Scheme 2

decarboxylases. The structure of SwMppP with D-Arg bound showed that the substrate is clamped in the active site with the guanidinium group of Arg352 interacting with the αcarboxylate group at one end and the carboxylates of Asp27 and Asp227 interacting with the guanidinium group at the other. The catalytic Lys221 makes a hydrogen bonding interaction with Asp218. Asp218 is an unusual addition to the phosphate-binding site of SwMppP and is shared only by the DOPA/aromatic L-amino acid decarboxylases. His29 of SwMppP appears to be unique to this class of enzyme and may be diagnostic for PLP-dependent hydroxylases. The involvement of Asp218 and His29 in the mechanism of SwMppPcatalyzed L-Arg hydroxylation is currently being investigated. This work allows us to propose an L-End biosynthetic scheme in which MppP converts L-Arg to 2-oxo-4-hydroxy-5guanidinovaleric acid, MppR cyclizes this to give the ketone analogue of L-End, and MppQ catalyzes a transamination to give the final product, L-End.

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aminotransferases (Figure S6), and the associated pairwise sequence identity matrix (Table S1). (PDF)

AUTHOR INFORMATION

Corresponding Author

*Address: 3210 N. Cramer St., Milwaukee, WI 53211. E-mail: [email protected]. Phone: 414-229-2647. Funding

This work was supported by Grant MCB-1157392 from the National Science Foundation, Directorate for Biological Sciences. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS Use of the Advanced Photon Source, an Office of Science User Facility operated for the U.S. Department of Energy (DOE) Office of Science by Argonne National Laboratory, was supported by the U.S. DOE under Contract DE-AC0206CH11357. Use of the LS-CAT Sector 21 was supported by the Michigan Economic Development Corp. and the Michigan Technology Tri-Corridor (Grant 085P1000817).

ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.5b01016. The full 1H NMR spectra for L-Arg (Figure S1) and LArg treated with SwMppP (Figure S2), 1H COSY spectrum of the L-Arg/SwMppP reaction mixture (Figure S3), the 1H/13C HSQC spectrum of the reaction mixture (Figure S4), the 1H/13C HMBC spectrum of the reaction mixture (Figure S5), a multiple sequence alignment of MppP-like enzymes, DOPA decarboxylases, and bacterial

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ABBREVIATIONS PLP, pyridoxal 5′-phosphate; PMP, pyridoxamine phosphate; LEnd, L-enduracididine; MPP, mannopeptimycin; ATase, aminotransferase; SwMppP, S. waddayamensis MppP.

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REFERENCES

(1) Ling, L. L., Schneider, T., Peoples, A. J., Spoering, A. L., Engels, I., Conlon, B. P., Mueller, A., Schaberle, T. F., Hughes, D. E., Epstein, S.,

J

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Biochemistry Jones, M., Lazarides, L., Steadman, V. A., Cohen, D. R., Felix, C. R., Fetterman, K. A., Millett, W. P., Nitti, A. G., Zullo, A. M., Chen, C., and Lewis, K. (2015) A new antibiotic kills pathogens without detectable resistance. Nature 517, 455−459. (2) Singh, M. P., Petersen, P. J., Weiss, W. J., Janso, J. E., Luckman, S. W., Lenoy, E. B., Bradford, P. A., Testa, R. T., and Greenstein, M. (2003) Mannopeptimycins, new cyclic glycopeptide antibiotics produced by Streptomyces hygroscopicus LL-AC98: antibacterial and mechanistic activities. Antimicrob. Agents Chemother. 47, 62−69. (3) Tsuchiya, K., and Takeuchi, Y. (1968) Enduracidin, an inhibitor of cell wall synthesis. J. Antibiot. 21, 426−428. (4) Ruzin, A., Singh, G., Severin, A., Yang, Y., Dushin, R. G., Sutherland, A. G., Minnick, A., Greenstein, M., May, M. K., Shlaes, D. M., and Bradford, P. A. (2004) Mechanism of action of the mannopeptimycins, a novel class of glycopeptide antibiotics active against vancomycin-resistant gram-positive bacteria. Antimicrob. Agents Chemother. 48, 728−738. (5) Kawakami, M., Nagai, Y., Fujii, T., and Mitsuhashi, S. (1971) Anti-microbial activities of enduracidin (enramycin) in vitro and in vivo. J. Antibiot. 24, 583−586. (6) Wong, C. T. T., Lam, H. Y., and Li, X. C. (2014) Effective synthesis of cyclic peptide yunnanin C and analogues via Ser/Thr ligation (STL)-mediated peptide cyclization. Tetrahedron 70, 7770− 7773. (7) Olivier, K. S., and Van Nieuwenhze, M. S. (2010) Synthetic studies toward the mannopeptimycins: synthesis of orthogonally protected beta-hydroxyenduracididines. Org. Lett. 12, 1680−1683. (8) Hatano, K., Nogami, I., Higashide, E., and Kishi, T. (1984) Biosynthesis of Enduracidin: Origin of Enduracididine and Other Amino Acids. Agric. Biol. Chem. 48, 1503−1508. (9) Magarvey, N. A., Haltli, B., He, M., Greenstein, M., and Hucul, J. A. (2006) Biosynthetic pathway for mannopeptimycins, lipoglycopeptide antibiotics active against drug-resistant gram-positive pathogens. Antimicrob. Agents Chemother. 50, 2167−2177. (10) Yin, X., and Zabriskie, T. M. (2006) The enduracidin biosynthetic gene cluster from Streptomyces fungicidicus. Microbiology 152, 2969−2983. (11) Goebel, N. C. (2012) Biosynthesis and Modification of the Antibiotic Enduracidin. School of Pharmacy, Oregon State University, Corvallis, OR. (12) Haltli, B., Tan, Y., Magarvey, N. A., Wagenaar, M., Yin, X., Greenstein, M., Hucul, J. A., and Zabriskie, T. M. (2005) Investigating beta-hydroxyenduracididine formation in the biosynthesis of the mannopeptimycins. Chem. Biol. 12, 1163−1168. (13) Burroughs, A. M., Hoppe, R. W., Goebel, N. C., Sayyed, B. H., Voegtline, T. J., Schwabacher, A. W., Zabriskie, T. M., and Silvaggi, N. R. (2013) Structural and functional characterization of MppR, an enduracididine biosynthetic enzyme from streptomyces hygroscopicus: functional diversity in the acetoacetate decarboxylase-like superfamily. Biochemistry 52, 4492−4506. (14) de Oliveira, L. G., Tormet Gonzalez, G. D., Samborsky, M., Marcon, J., Araujo, W. L., and de Azevedo, J. L. (2014) Genome Sequence of Streptomyces wadayamensis Strain A23, an Endophytic Actinobacterium from Citrus reticulata. Genome Announcements 2, e00625-14. (15) Otwinowski, Z., and Minor, W. (1997) Processing of X-ray Diffraction Data Collected in Oscillation Mode. Methods Enzymol. 276, 307−326. (16) Vonrhein, C., Blanc, E., Roversi, P., and Bricogne, G. (2006) Automated structure solution with autoSHARP. Methods Mol. Biol. 364, 215−230. (17) Cowtan, K. (2006) The Buccaneer software for automated model building. 1. Tracing protein chains. Acta Crystallogr., Sect. D: Biol. Crystallogr. 62, 1002−1011. (18) McCoy, A. J., Grosse-Kunstleve, R. W., Adams, P. D., Winn, M. D., Storoni, L. C., and Read, R. J. (2007) Phaser crystallographic software. J. Appl. Crystallogr. 40, 658−674.

(19) Emsley, P., Lohkamp, B., Scott, W. G., and Cowtan, K. (2010) Features and development of Coot. Acta Crystallogr., Sect. D: Biol. Crystallogr. 66, 486−501. (20) Afonine, P. V., Mustyakimov, M., Grosse-Kunstleve, R. W., Moriarty, N. W., Langan, P., and Adams, P. D. (2010) Joint X-ray and neutron refinement with phenix.refine. Acta Crystallogr., Sect. D: Biol. Crystallogr. 66, 1153−1163. (21) Word, J. M., Lovell, S. C., Richardson, J. S., and Richardson, D. C. (1999) Asparagine and glutamine: using hydrogen atom contacts in the choice of side-chain amide orientation. J. Mol. Biol. 285, 1735− 1747. (22) Chen, V. B., Arendall, W. B., 3rd, Headd, J. J., Keedy, D. A., Immormino, R. M., Kapral, G. J., Murray, L. W., Richardson, J. S., and Richardson, D. C. (2010) MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr., Sect. D: Biol. Crystallogr. 66, 12−21. (23) Urzhumtseva, L., Afonine, P. V., Adams, P. D., and Urzhumtsev, A. (2009) Crystallographic model quality at a glance. Acta Crystallogr., Sect. D: Biol. Crystallogr. 65, 297−300. (24) Moriarty, N. W., Grosse-Kunstleve, R. W., and Adams, P. D. (2009) electronic Ligand Builder and Optimization Workbench (eLBOW): a tool for ligand coordinate and restraint generation. Acta Crystallogr., Sect. D: Biol. Crystallogr. 65, 1074−1080. (25) Goldberg, J. M., and Kirsch, J. F. (1996) The reaction catalyzed by Escherichia coli aspartate aminotransferase has multiple partially rate-determining steps, while that catalyzed by the Y225F mutant is dominated by ketimine hydrolysis. Biochemistry 35, 5280−5291. (26) Lu, Z., Nagata, S., McPhie, P., and Miles, E. W. (1993) Lysine 87 in the beta subunit of tryptophan synthase that forms an internal aldimine with pyridoxal phosphate serves critical roles in transimination, catalysis, and product release. J. Biol. Chem. 268, 8727−8734. (27) Sun, X., Li, H., Alfermann, J., Mootz, H. D., and Yang, H. (2014) Kinetics profiling of gramicidin S synthetase A, a member of nonribosomal peptide synthetases. Biochemistry 53, 7983−7989. (28) Bertoldi, M., and Borri Voltattorni, C. (2000) Reaction of dopa decarboxylase with L-aromatic amino acids under aerobic and anaerobic conditions. Biochem. J. 352 (Part2), 533−538. (29) Bertoldi, M., and Borri Voltattorni, C. (2003) Reaction and substrate specificity of recombinant pig kidney Dopa decarboxylase under aerobic and anaerobic conditions. Biochim. Biophys. Acta, Proteins Proteomics 1647, 42−47. (30) Hayashi, H., Tsukiyama, F., Ishii, S., Mizuguchi, H., and Kagamiyama, H. (1999) Acid-base chemistry of the reaction of aromatic L-amino acid decarboxylase and dopa analyzed by transient and steady-state kinetics: preferential binding of the substrate with its amino group unprotonated. Biochemistry 38, 15615−15622. (31) Bunik, V. I., Schloss, J. V., Pinto, J. T., Dudareva, N., and Cooper, A. J. (2011) A survey of oxidative paracatalytic reactions catalyzed by enzymes that generate carbanionic intermediates: implications for ROS production, cancer etiology, and neurodegenerative diseases. Adv. Enzymol. Relat. Areas Mol. Biol. 77, 307− 360. (32) Mehta, P. K., and Christen, P. (1993) Homology of pyridoxal5′-phosphate-dependent aminotransferases with the cobC (cobalamin synthesis), nifS (nitrogen fixation), pabC (p-aminobenzoate synthesis) and malY (abolishing endogenous induction of the maltose system) gene products. Eur. J. Biochem. 211, 373−376. (33) Okamoto, A., Higuchi, T., Hirotsu, K., Kuramitsu, S., and Kagamiyama, H. (1994) X-ray crystallographic study of pyridoxal 5′phosphate-type aspartate aminotransferases from Escherichia coli in open and closed form. J. Biochem 116, 95−107. (34) Krissinel, E., and Henrick, K. (2004) Secondary-structure matching (SSM), a new tool for fast protein structure alignment in three dimensions. Acta Crystallogr., Sect. D: Biol. Crystallogr. 60, 2256− 2268. (35) Eliot, A. C., and Kirsch, J. F. (2004) Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations. Annu. Rev. Biochem. 73, 383−415. K

DOI: 10.1021/acs.biochem.5b01016 Biochemistry XXXX, XXX, XXX−XXX

Article

Biochemistry

by site-directed mutagenesis and kinetic studies. Biochemistry 44, 13970−13980. (54) Giardina, G., Montioli, R., Gianni, S., Cellini, B., Paiardini, A., Voltattorni, C. B., and Cutruzzola, F. (2011) Open conformation of human DOPA decarboxylase reveals the mechanism of PLP addition to Group II decarboxylases. Proc. Natl. Acad. Sci. U. S. A. 108, 20514− 20519. (55) Burkhard, P., Dominici, P., Borri-Voltattorni, C., Jansonius, J. N., and Malashkevich, V. N. (2001) Structural insight into Parkinson’s disease treatment from drug-inhibited DOPA decarboxylase. Nat. Struct. Biol. 8, 963−967. (56) Bertoldi, M., Cellini, B., Montioli, R., and Borri Voltattorni, C. (2008) Insights into the mechanism of oxidative deamination catalyzed by DOPA decarboxylase. Biochemistry 47, 7187−7195. (57) Fenn, T. D., Ringe, D., and Petsko, G. A. (2003) POVScript+: a program for model and data visualization using persistence of vision ray-tracing. J. Appl. Crystallogr. 36, 944−947. (58) Kraulis, P. (1991) MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J. Appl. Crystallogr. 24, 946−950.

(36) Schneider, G., Kack, H., and Lindqvist, Y. (2000) The manifold of vitamin B6 dependent enzymes. Structure 8, R1−6. (37) Battchikova, N., Koivulehto, M., Denesyuk, A., Ptitsyn, L., Boretsky, Y., Hellman, J., and Korpela, T. (1996) Aspartate aminotransferase from an alkalophilic Bacillus contains an additional 20-amino acid extension at its functionally important N-terminus. J. Biochem. 120, 425−432. (38) McPhalen, C. A., Vincent, M. G., and Jansonius, J. N. (1992) Xray structure refinement and comparison of three forms of mitochondrial aspartate aminotransferase. J. Mol. Biol. 225, 495−517. (39) Wrenger, C., Muller, I. B., Schifferdecker, A. J., Jain, R., Jordanova, R., and Groves, M. R. (2011) Specific inhibition of the aspartate aminotransferase of Plasmodium falciparum. J. Mol. Biol. 405, 956−971. (40) Kirsch, J. F., Eichele, G., Ford, G. C., Vincent, M. G., Jansonius, J. N., Gehring, H., and Christen, P. (1984) Mechanism of action of aspartate aminotransferase proposed on the basis of its spatial structure. J. Mol. Biol. 174, 497−525. (41) Yano, T., Kuramitsu, S., Tanase, S., Morino, Y., and Kagamiyama, H. (1992) Role of Asp222 in the catalytic mechanism of Escherichia coli aspartate aminotransferase: the amino acid residue which enhances the function of the enzyme-bound coenzyme pyridoxal 5′-phosphate. Biochemistry 31, 5878−5887. (42) Yano, T., Mizuno, T., and Kagamiyama, H. (1993) A hydrogenbonding network modulating enzyme function: asparagine-194 and tyrosine-225 of Escherichia coli aspartate aminotransferase. Biochemistry 32, 1810−1815. (43) Krissinel, E., and Henrick, K. (2004) Secondary-structure matching (SSM), a new tool for fast protein structure alignment in three dimensions. Acta Crystallogr., Sect. D: Biol. Crystallogr. 60, 2256− 2268. (44) Krissinel, E., and Henrick, K. (2005) Multiple Alignment of Protein Structures in Three Dimensions. In Computational Life Sciences (Berthold, M., Glen, R., Diederichs, K., Kohlbacher, O., and Fischer, I., Eds.) pp 67−78, Springer, Berlin. (45) Krupka, H. I., Huber, R., Holt, S. C., and Clausen, T. (2000) Crystal structure of cystalysin from Treponema denticola: a pyridoxal 5′-phosphate-dependent protein acting as a haemolytic enzyme. EMBO J. 19, 3168−3178. (46) Clausen, T., Schlegel, A., Peist, R., Schneider, E., Steegborn, C., Chang, Y. S., Haase, A., Bourenkov, G. P., Bartunik, H. D., and Boos, W. (2000) X-ray structure of MalY from Escherichia coli: a pyridoxal 5′-phosphate-dependent enzyme acting as a modulator in mal gene expression. EMBO J. 19, 831−842. (47) Goto, M., Omi, R., Miyahara, I., Hosono, A., Mizuguchi, H., Hayashi, H., Kagamiyama, H., and Hirotsu, K. (2004) Crystal structures of glutamine:phenylpyruvate aminotransferase from Thermus thermophilus HB8: induced fit and substrate recognition. J. Biol. Chem. 279, 16518−16525. (48) Chu, L., Ebersole, J. L., Kurzban, G. P., and Holt, S. C. (1997) Cystalysin, a 46-kilodalton cysteine desulfhydrase from Treponema denticola, with hemolytic and hemoxidative activities. Infect. Immun. 65, 3231−3238. (49) Chu, L., and Holt, S. C. (1994) Purification and characterization of a 45 kDa hemolysin from Treponema denticola ATCC 35404. Microb. Pathog. 16, 197−212. (50) Zdych, E., Peist, R., Reidl, J., and Boos, W. (1995) MalY of Escherichia coli is an enzyme with the activity of a beta C-S lyase (cystathionase). J. Bacteriol. 177, 5035−5039. (51) Hosono, A., Mizuguchi, H., Hayashi, H., Goto, M., Miyahara, I., Hirotsu, K., and Kagamiyama, H. (2003) Glutamine:phenylpyruvate aminotransferase from an extremely thermophilic bacterium, Thermus thermophilus HB8. J. Biochem. 134, 843−851. (52) Kezuka, Y., Yoshida, Y., and Nonaka, T. (2012) Structural insights into catalysis by betaC-S lyase from Streptococcus anginosus. Proteins: Struct., Funct., Genet. 80, 2447−2458. (53) Cellini, B., Bertoldi, M., Montioli, R., and Borri Voltattorni, C. (2005) Probing the role of Tyr 64 of Treponema denticola cystalysin L

DOI: 10.1021/acs.biochem.5b01016 Biochemistry XXXX, XXX, XXX−XXX